Preparation and use of biofilm-degrading, multiple-specificity, hydrolytic enzyme mixtures

ABSTRACT

The present invention relates to isolated structures containing degradative enzymes produced from a marine organism. The enzymes produced are based on the carbon source upon which the marine organism is growing. The enzymes are found in structures that can be isolated such that the degradative enzymes are easily harvested.

FIELD OF THE INVENTION

The present invention relates to a method for preparing biofilmdegrading, multiple specificity, hydrolytic enzyme mixtures which arespecifically tailored to remove targeted biofilms.

The present invention also is directed to methods for using hydrolyticenzyme mixtures in both industrial and therapeutic applications. Theindustrial applications include but are not limited to the use ofbiofilm-degrading, multiple specificity, hydrolytic enzyme mixtures forremoving or preventing the formation of biofilms in water coolingtowers, industrial process piping, heat exchangers, in food processingor food preparation, in potable water systems, reservoirs, swimmingpools, or related sanitary water systems, and on membranes such as thoseused for desalinization, industrial processes, or related applications.

The therapeutic applications include but are not limited to the use oftherapeutically-useful, multiple-specificity, hydrolytic enzyme mixturesor components thereof for the prevention or treatment of dental cariesand periodontal disease, improving the treatment of cystic fibrosis orthe complications or symptoms of cystic fibrosis, and diseases orcomplications associated with biofilm formation on implantable medicaldevices such as cardiovascular devices. The route of administration canbe by any means including delivering the hydrolytic enzyme mixture byaerosol to the lungs and applying the hydrolytic enzyme mixturetopically.

BACKGROUND OF THE INVENTION

Naturally occurring biofilms are continuously produced and oftenaccumulate on numerous industrial surfaces and on biological surfaces.In an industrial setting, the presence of these biofilms causes adecrease in the efficiency of industrial machinery, requires increasedmaintenance, and presents potential health hazards. For example, thesurfaces of water cooling towers become increasingly coated withmicrobially produced biofilm slime which both constricts water flow andreduces heat exchange capacity. Water cooling tower biofilms may alsoharbor pathogenic microorganisms such as Legionella pneumophila. Foodpreparation lines are routinely plagued by biofilm build-up both on themachinery and on the food product where biofilms often include potentialpathogens. Industrial biofilms are complex assemblages of insolublepolysaccharide-rich biopolymers which are produced and elaborated bysurface dwelling microorganisms. The chemical composition of industrialbiofilms are diverse and are specific to each species of surfacedwelling microorganism. Because of this complexity and diversity,non-specific hydrolytic enzymes are ineffective in degrading thesebiofilms and consequently ineffective in reducing or eliminating theundesirable biofilm.

On a biological surface, the presence of these biofilms results in thegrowth of, and subsequent colonization by, pathogenic microorganisms onan internal or external surface of a host animal or on the surface ofobjects introduced into the animal (e.g surgical implants). Animalpathogens which colonize surfaces are often maintained and protected byunique polysaccharide rich biofilms produced by the pathogen. Suchbiofilms coat the infected or colonized surface of the animal orimplanted object and continue to be produced during the disease process.For many diseases, biofilms are required for the disease process tobecome established and to progress. The chemical compositions ofpathogen-associated surface biofilms, which consist of complex mixturesof biopolymers, are specific to each species of pathogen. Because ofthis complexity, non-specific hydrolytic enzymes or hydrolytic enzymeswith a single specificity are ineffective in degrading these biofilmsand consequently ineffective in reducing or eliminating the diseasecondition. At the present time, there are no therapeutic products whichare commercially employed to degrade and remove these disease related,pathogen-produced biofilms.

Currently, biofilms are most commonly removed using physical abrasion, aprocess which is both inefficient and incomplete. Antimicrobials(biocides and antibiotics) are employed to slow biofilm build-up bykilling the microbes that produce biofilms; however, once established,the biofilms protect the embedded, biofilm-producing bacteria from theaction of these agents. Furthermore, many antimicrobial agents are toxicand damaging to the environment. Consequently, there is a need for amethod to readily remove and control biofilms that does not dependsolely on physical abrasion or on the action of antimicrobial agents.This need could be met by a mixture of multiple specificity, hydrolyticenzymes which have been tailored to degrade the specific complexbiopolymer composition of a target biofilm. A tailored mixture ofmultiple hydrolytic enzymes could be employed to degrade biofilmsresulting in their more complete removal and in enhanced antimicrobialactivity.

It has recently become apparent that insoluble complex polysaccharides(ICP) in the environment are most efficiently degraded by a cascade ofenzymes acting in concert. The degradation of these insoluble complexpolysaccharides require more than “simple” exoenzymes. Normally, anarray of enzymes, part of a complex system, is required to fullyhydrolyze the polysaccharide into its final monosaccharide end product(Belas et al., 1988; Bassler et al., 1991 b; Bayer & Lamed 1992; Salyerset al., 1996; Svitil et al., 1997). Most of the carbohydrate-degradingenzymes are highly specific for glycosidic sugar and the anomericconfiguration of the glycosidic bond.

They can act endolytically, hydrolyzing internal carbohydrate bonds,generating oligosaccharide intermediates resulting in relatively rapidviscosity decreases of the polymer; others act exolytically, degradingthe polymer from the non-reducing termini generating a singlemonosaccharide end product.

These enzymes tend to show a higher specificity with high molecularweight substrates than lower molecular weight substrates.

For the degradation of the insoluble complex polysaccharides, enzymelocalization relative to other enzymes and biomolecules is oftenimportant for enzyme efficiency, as is the chemistry of its active site.Many reports have been published describing the properties of numerousisolated polysaccharide-degrading bacteria; however, relatively littleis understood concerning how intact bacteria degrade insoluble complexpolysaccharides or how the multiple enzymes produced by the organisminteract (Salyers et al., 1996). It should be noted that degradation ofthe insoluble complex polysaccharides into its monosaccharide requiresmultiple enzymes and possibly other proteins (e.g. substrate-binding).

The present invention teaches general methods for preparingbiofilm-degrading, multiple-specificity hydrolytic enzyme mixtures whichare specifically tailored to remove targeted industrial and/ordisease-related biofilms. These biofilm degrading hydrolytic enzymemixtures can be employed to remove or degrade biofilms from the targetsurface causing a reduction of the biofilm and resulting in increasedefficiency and improved hygiene in industrial settings and in improvedtreatment in therapeutic settings. The present invention will findapplication in numerous settings where biofilms currently presentefficiency and health problems.

Hydrolytic enzyme mixtures can be employed, via direct application tothe biofilm, to remove or degrade disease-associated and/or industrialbiofilms from the surfaces colonized by the pathogen. The presentinvention will find application in industrial settings, such as watercooling towers, waste water piping, heat exchangers, and foodpreparation lines. The present invention will also find application as atherapeutic agent for the treatment of numerous currently uncontrolledanimal, and particularly human, diseases. For example: i) Oralplaque-forming bacterial species, the causal agents of dental caries,are maintained by complex biofilms required for their continuedcolonization of the tooth surface and their disease causing action.Animal species, particularly humans, exposed to these oralplaque-forming bacteria are at risk of developing caries. Thesepathogen-related biofilms are currently removed by physical abrasion.ii) Porphyromonas gingivalis, the causal agent of periodontal disease,requires a glycocalyx biofilm for its disease action. Human periodontaldisease is currently the major cause of tooth loss world-wide. iii)Cystic fibrosis, which has a frequency of 1 in every 2,000 live births,frequently is associated with infection by Pseudomonas aeruginosa in thelungs, P. aeruginosa produces a complex, alginate-based biofilm whichdirectly results in the hyperviscous mucus characteristic of cysticfibrosis patients. This biofilm is also the substrate for pulmonaryinfections by opportunistic pathogens characteristic of the disease. iv)Implantable medical devices, such as artificial valves, stents, andcatheters, can become colonized by pathogens such as Streptococcus sp.,leading to premature failure of the devices and/or life-threateningsecondary infections. v) contact lenses can become coated with biofilmsand colonized by pathogens. The enzyme mixtures of the present inventionwill conveniently remove these biofilms.

Microorganisms which degrade complex polysaccharides are known in theart. Some marine microorganisms faced with oligotrophic conditions inthe pelagic zone, have evolved powerful enzyme systems to take advantageof the ubiquitous marine snow, which are potential oases in thenutritionally poor open waters. As a consequence, selected marinespecies have developed very efficient mechanisms to utilize complexpolysaccharides. Marine bacterium Microbulbifer [e.g. 2-40 (deposited atthe American Type Culture Collection as ATCC 43961) and IRE-31(deposited at the American Type Culture Collection as ATCC 700072)] andMarinobacterium [e.g. KW-40 (deposited at the American Type CultureCollection as ATCC 700074)] have been identified as a potentiallyimportant bioremediation species, since they synthesize an unusuallylarge number of degradative enzymes. Marine bacterium Microbulbifer 2-40is described in U.S. Pat. No. 5,418,156 (described as Alteromonas 2-40in U.S. Pat. No. 5,418,156 but subsequently determined through nucleicacid sequencing to be a Microbulbifer) which is hereby incorporated byreference into the present document. The marine/estuarine bacterium,2-40, is a periphytic organism isolated from a salt marsh growing onSpartina alterniflora. It is Gram negative, pleomorphic, rod-shaped andmotile. This aerobe requires sea salts and carbohydrates for growth. Itproduces numerous proteases, lipases, and carbohydrases that allowMicrobulbifer to degrade a variety of complex, insoluble polysaccharidesof plant, fungi, and animal origin. These polysaccharides includealginate, araban, carrageenan, carboxymethylcellulose, chitin, glycogen,β-glucan, pectin, laminarin, pullulan, starch, xylan, and agar.

Relatively recently, a novel structure relating to insoluble substratedegradation was discovered in a Gram positive bacterium. It was acellulose-binding and multicellulase-containing cell-surfaceprotuberance produced by Clostridium thermocellum and C. cellulovorans.Coined “cellulosomes” they were found to attach directly to theinsoluble substrates, via special cellulose binding proteins. Thus, theybring cellulases into contact with cellulose, targeting the enzymesubstrate complex. Cellulosomes are comprised of at least 14 differentproteins. Cip A refers to the largest of the cellulosome proteins,approximately 250 kDa. It serves as the scaffolding protein, binding andanchoring the enzymatic components and securing the entire cellulosomeon the cell surface. This protein has a 166 amino acid sequence that isrepeated 9 times and is a receptor for the enzymatic cellulosomecomponents, such as CeID. CeID is a 68 kDa endoglucanase isolated fromthe cellulosome whose carboxy-terminus has a docking sequence that bindsto the CipA receptors.

Such structures (hereinafter “degradosomes”) may be involved in thedepolymerization of other insoluble polymers in addition to cellulose.Degradosome components could also consist of spatially arrayed enzymes,adhesions and scaffold protein. Not only do degradosomes maintain thereleased monomer product close to the cell for metabolic utilization,but the degradosome may place a cascade of hydrolytic enzymes in properjuxtaposition for optimal enzyme activity. It is also an attachmentorganelle, incorporating specific polymer binding proteins. Because ofwhole cell/degradosome efficiency and the potential for continued enzymesynthesis, the use of living bacteria as bioreactors in the degradationof not only cellulose, but also potential chitin (aquaculture), algaeslimes (algal culture) and biofouled surfaces may be quite advantageous.

It has been found that Microbulbifer expresses cell surfaceprotruberances on its outer membrane and that they are expressedcoincidentally with insoluble biopolymer degradation. Furthermore,results suggest that insoluble carbohydrate degradation is indeed mostefficient in Microbulbifer when the carbohydrase systems are introducedand degradosome structures are expressed on the outer membrane of thisGram negative rod. Microbulbifer has been shown to a) synthesize greaterquantities, and a greater variety, of degradable carbohydrase systemswhen grown in media containing several complex carbohydrate carbonsources than when grown in a single complex carbohydrate minimal media,b) package agarases and chitinases in different degradosome structuresfrom one another, and c) undergo morphogenesis and synthesizeinteresting tubular is structures under conditions of carbon limitation.In Microbulbifer a system of enzymes in the degradosome, acting inconcert, degrade a portion of the carbohydrate to monomers, thusconverting waste into usable nutrients.

Living Microbulbifer may be used for bioremediation since it not apathogen of animals or invertebrates. Other genera shown to synthesizepolysaccharide degrading enzymes (e.g. agarases) include Vibrio,Alteromonas, Flavobacterium, Streptomyces, and Pseudomonas.Microbulbifer produces three agarases with activities which areanalogous to those of P. atlantica. However, the Microbulbifer agaraseshave different molecular weights, higher specific activity and aregenerally more resistant to denaturation than those of other species.

Alginate is commonly produced by both algae, such as Macrocystispyrifera, and prokaryotes, such as Azotobacter vinelandii, and isconsequently a major component of many biofilms. The alginic acid ofmucoid Pseudomonas aerugnosa is of medical importance in theexacerbation of cystic fibrosis where it acts as a virulence factor,inhibiting host phagocytosis. Bacterial alginates differ from algalalginates in the degree of O-acetylation of the mannuronic acidresidues. Chronic pulmonary infection with Pseudomonas aeruginosa is amajor cause of mortality in cystic fibrosis patients. Pseudomonasaeruginosa produces a number of virulence factors includingextracellular toxins, proteases, hemolysins and exopolysaccharides. Theexopolysaccharide alginate shields the bacterium from the host defensemechanisms and anti-microbial agents. The exopolysaccharides may alsopromote adherence of mucoid strains to the epithelial cells of therespiratory tract. The use of an alginate lyase obtained fromFlavobactedum OTC-6 as a therapeutic medicine for cystic fibrosis isdescribed in U.S. Pat. No. 5,582,825. The alginate enzyme systemproduced by Microbulbifer differs from alginases purified from otherorganisms in that Microbulbifer produces an enzyme system made up ofseveral enzymes which act together to more effectively degradepolysaccharides.

Pseudomonas aeruginosa infections also occur in burn victims,individuals with cancer and patients requiring extensive stays inintensive care units. Therefore, these patients would also benefit froman improved method for treating Pseudomonas aeruginosa infections.

In addition, many strains of Streptococcus mutans have been shown to becariogenic in experimental animals and are directly associated withhuman dental caries (Hardie, J, M., 1981. The microbiology of dentalcaries. In: Silverstone, Johson, Hardie and Williams (ed.), DentalCaries: Aetiology, Pathology, and Prevention, The Macmillian Press Ltd,London, pp 48-69.; Tanzer, J. M. (ed), 1981, Animal Models in Cariology,Special Supplement, Microbiology Abstracts, Information Retrieval,Washington D.C. and London.) and can be isolated from cases of infectiveendocarditis. The primary habitat of S. mutans is the tooth surface ofhumans, and its colonization of this surface is favored by high levelsof dietary sucrose. S. mutans produces biofilms which are composed ofseveral types of extracellular polysaccha rides which are manufacturedfrom sucrose and which are important in the colonization of hard tissuesurfaces in the mouth (Gibbons, R. J. and J. van Houte, 1973. On theformation of dental plaques. J. Periodontal. 44-347-360.; Gibbons, R. J.and J. van Houte, 1975. Bacterial adherence in oral microbial ecology.Ann. Rev. Microbiol. 29:19-44.; Hamada, S. and H. D. Slade, 1980.Mechanisms of adherence of Streptococcus mutans to smooth surfaces invitro. In: Beachey (ed.), Bacterial Adherence, Chapman and Hall, London,pp. 105-135.) These glucans include a water soluble α-(1-6)-linkedlinear glucose polymer with α(1-3) glucosidic branch linkages (Long, L.and J. Edwards, 1972. Detailed structure of a dextran from a cariogenicbacterium. Carbohydr. Res. 24:216-217.), and other essentiallywater-insoluble, cell-associated polymers. These water-insolublepolymers contain a high proportion of α(1-3) glucosidic linkages and aregenerally resistant to degradation by enzymes commonly present in theoral cavity. Because these S. mutans produced biofilms are resistant toenzymatic degradation they build up on the tooth surface, are a majorcomponent of dental plaque, and provide an additional habitat for dentalcary causing microbes and microbes which contribute to “bad breath.”Currently, dental plaque is removed by physical scraping of the toothsurface which is most often performed by dental technicians. The S.mutans biofilm is only partially removed from the tooth surface bybrushing with a dentifrice or by mouthwash. Consequently, an enzymaticpreparation able to degrade the S. mutans produced polysaccharidebiofilm and aid in the removal of dental plaque could be incorporatedinto a toothpaste, into a mouth rinse, or into other vehicles whichcontact the tooth surface. Enzymatically degraded S. mutans biofilm andthe biofilm-associated microorganisms can be more easily and readilyremoved from the oral cavity resulting in fewer dental caries andobjectionable mouth odors.

In view of the above discussion, one object of the present invention isto develop nontoxic, environmentally friendly methods for removingindustrial biofilms.

Another object of the present invention is to develop a method forremoving various disease related biofilms on an internal or externalsurface of an animal or on an implant prior to or after implantation inan animal by applying to the affected surface or administering to theanimal an effective amount of a) an organism which produces a hydrolyticenzyme mixture, b) a hydrolytic enzyme mixture and/or c) a component ofa hydrolytic enzyme mixture.

SUMMARY OF THE INVENTION

In the present invention, multiple specificity, hydrolytic enzymemixtures are produced using certain bacterial species (e.g. marinesaprophytic bacteria such as Microbulbifer 2-40 also known asAlteromonas 2-40). The bacteria are selected for their ability to growon and catabolize or degrade a wide range of complex polysaccharidesources such as those that are present in biofilms. The bacteria arecultured in the presence of one or more polysaccharides which arepresent in the targeted biofilm. The polysaccharides are used as theprimary carbon source to support the growth and metabolism of thebacteria. During the growth of the bacteria in the presence of thepolysaccharide, a mixture of hydrolytic enzymes with multiplespecificities capable of degrading a complex biofilm material containingthe polysaccharide is produced. By altering the composition of thepolysaccharides in the culture medium, a custom tailored mixture ofenzymes can be produced. The hydrolytic enzyme mixture can then beisolated and applied to the affected industrial or biological surface.

The industrial application of a multiple specificity, hydrolytic enzymemixture will remove or degrade biofilms from the target industrialsurface causing a reduction of the biofilm thereby resulting inincreased efficiency and improved hygiene.

The therapeutic administration of a hydrolytic enzyme mixture or acomponent thereof will reduce the biofilm and thereby enable antibioticsand/or the animal recipient's immune system to fight an infection with abacterial pathogen. The therapeutic, multiple specificity, hydrolyticenzyme mixture of the present invention will therefore be useful as anadjunct to standard anti-infective therapies when a biofilm producingpathogen is involved.

When used therapeutically, the hydrolytic enzyme mixture of the presentinvention can be administered by any route, including but not limited tooral, pulmonary (by aerosol or by other respiratory device forrespiratory tract infections), nasal, IV, IP and intra-ocularly.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1. Growth in monosaccharides or ICP as sole carbon sources. 2-40was grown in MM+0.2% of the indicated sole carbon source. Batch culturegrowth (OD_(600nm)) was plotted vs. time: Growth in monosaccharides (A.)and Growth in ICP (B. & C.). Growth in other sole carbon sources isshown in subsequent figures: glucose, FIG. 2; agarose, FIG. 3; andchitin, FIG. 4. The cultures were sampled in time course for enzymeactivity (see Table 4).

FIG. 2. Batch growth in 0.2% glucose minimal medium. A.

Batch culture growth (Abs 600 nm)(-∘-) and cell number (cfu/ml)(-□-)were plotted vs time. Inset graph shows the relationship between OD(600nm)(y-axis) and cell number (cfu/ml)(x-axis) in glucose MM vs a glucoseMM blank (-∘-). Solid black line on inset graph is the linearregression.

FIG. 3. Batch growth in 0.2% agarose minimal medium. A. Batch culturegrowth (Abs 600 nm) (-Δ-) and cell number (cfu/ml) (-∘-) were plotted vstime. Inset graph shows the relationship between OD (600 nm)(y-axis) andcell number (cfu/ml)(x-axis) in agarose MM (-∘-). Solid line on theinset graph is the linear regression.

FIG. 4. Batch growth in 0.2% chitin minimal medium. A.

Batch culture growth (Abs 600 nm)(-∘-) and relative extracellularchitinase activity (-□-)(μg/ml reducing sugar) were plotted vs time. B.Relative carbohydrase activity (μg/ml reducing sugar) of cellularchitinases (-□-) and cellular agarases (-Δ-) produced by 2-40 grown inchitin MM (graphed in A.) were plotted vs time. There was no detectableextracellular agarase activity in 2-40 grown in chitin MM.

FIG. 5. Fine structure of 2-40 grown in 0.2% glucose (A), agarose (B),or chitin (C). Cells grown in the respective substrate to mid-log phasewere observed by SEM. Note the rough surface (degradosome covered) ofcells grown in ICP (B & C) and their absence in glucose (A). Theproduction of these structures correlated to the respective, homologousICP degradase activity. Scale bar=300 nm.

FIG. 6. Immunolabeling of agarase and chitinase synthesizedsimultaneously by 2-40. Whole cells of 2-40, grown to mid-log phase inMM containing 0.2% agarose and chitin, were labeled with anti-agaraseantibody (primary) and 10 nm gold labeled goat anti-rabbit IgG antibody(secondary). They were additionally labeled with 20 mm gold labeledanti-chitinase antibody. Cells were stained with 1% uranyl acetate andobserved by TEM. Arrows indicate agarase labeling. Note the segregationof agarase (small gold particles) and chitinase (larger gold isparticles) labeling in the micrograph.

FIG. 7. Zymogram of 8% native-PA gel overlay with molecular weightsadded for reference. For both A and B Lane 1, 60 μg total protein of0.2% glucose grown 2-40; Lane 2, 40 μg total protein of partiallypurified concentrated alginase preparations. Duplicate lanes of glucose2-40 cell prep and alginase prep from the gel were (A) silver stainedand (B) used in the zymogram overlay, then stained with toluidine blueO. There are eight bands with alginase activity with approximatemolecular weight of 87, 66, 43, 39, 35, 27, 25 and 23 kD.

FIG. 8. Sequence from position 300 to 1500 of the 16S gene ofMicrobulbifer 240.

DETAILED DESCRIPTION OF THE INVENTION

The general methods and steps for preparing biofilm-degrading,multiple-specificity, hydrolytic enzyme mixtures are as follows.

First, certain bacterial species (e.g. marine saprophytic bacteria asexemplified by Microbulbifer) are selected for their ability to grow onand catabolize or degrade a wide range of complex polysaccharide sourcesincluding those that comprise biofilms. These selected bacterial speciesare then cultured in a medium or series of media containing one or moreof the specific polysaccharides comprising the targeted biofilm, orderivatives thereof. The polysaccharides are used as the primary carbonsource to support the growth and metabolism of the bacterial species.During growth of the bacterial species on this specialized medium, amixture of hydrolytic enzymes with multiple specificities capable ofdegrading the complex biofilm material is produced on the surface of theorganism in enzyme containing protuberances [for example through theformation of enzyme-containing appendages (degradosomes) from thebacterial cell surface] and elaborated from the cells in tubules orvesicles or otherwise released into the medium by the bacteria inincreasing quantities as the insoluble complex polysaccharides aredepleted, as exemplified by certain marine saprophytic bacteria such asMicrobulbifer 2-40. By altering the composition of the polysaccharidesor their derivatives in the culture media, a custom-tailored consortiaof hydrolytic enzymes can be produced. The biofilm-degrading,multiple-specificity, hydrolytic enzyme mixtures are separated from theculture, preferably from the culture supernatant and more preferablyfrom supernatant having hydrolytic enzyme-containing appendages orvesicles. The enriched hydrolytic enzyme mixture is appropriatelyformulated and applied to the biofilm which results in the degradationand removal of the biofilm targeted for the application. Alternatively,the living organism itself, the degradosomes, tubules, vesicles orpurified enzymes can be directly applied to the biofilm. It should beclear that since each biofilm forming microbial species produces aunique biofilm material, each biofilm will require a different, customtailored, multiple-specificity, hydrolytic enzyme mixture to achievebiofilm control. These different mixtures can be produced and tailoredfor each use by employing the targeted biofilm material as the primarycarbon source during the culture of the bacterial species. The enzymemixture can then be purified and applied to the targeted biofilm.

The enzyme mixture can be purified as a mixture or the various enzymesystems present in the mixture can be purified individually. If theenzymes are purified individually an enzyme mixture can be reformulatedafter purification or the enzymes can be used individually (e.g. in somespecific therapeutic applications). It is not necessary to completelypurify the enzyme systems prior to use. The enzymes may be present inand purified from degradosomes, vesicles or tubules or the degradosomes,vesicles or tubules can be applied directly to the targeted biofilm.

In an industrial setting the amount of the enzyme mixture to be appliedto the targeted biofilm is not critical. The amount to be applied caneasily be determined by routine experimentation and will be related tothe composition of the biofilm. In an industrial setting, the enzymemixture is applied by contacting the targeted biofilm with theappropriate enzyme mixture.

In a therapeutic application, there is no particular limitation on themodality of treatment with the enzyme mixture of this invention and thecomposition can be administered according to a treatment protocol whichdepends on the patient's age, sex and other factors, the severity ofdisease, etc. A spray or an infusion can be directly applied to theaffected site. A tablet, solution, emulsion, powder or capsule can beadministered orally. An injection can be administered in admixture withan ordinary infusion fluid such as glucose solution, amino acidinfusion, etc. Thus, the routes of administration include but are notlimited to: oral, aerosol or other device for delivery to the lungs,nasal spray, intravenous, intramuscular, intraperitoneal, and topical.Excipients which can be used as a vehicle for the delivery of the enzymemixture will be apparent to those skilled in the art. For example, theenzyme mixture could be in lyophilized form and be dissolved just priorto administration or the enzyme mixture could be present in liposomes.If the targeted biofilm is on an oral surface, the enzyme mixture couldbe applied in the form of a toothpaste or mouth rinse. The dosage ofadministration of the enzyme mixture for reducing biofilms on an oralsurface is between 0.1 mg-1 g per ml of delivery excipient. The dosageof administration of the enzyme mixture for treating P. aeruginosainfections is contemplated to be in the range of about 0.1-100 mg/per kgbody weight, and preferably about 1-10 mg/per kg body weight.

With respect to the aerosol administration to the lungs, the hydrolyticenzyme mixture is incorporated into an aerosol formulation specificallydesigned for administration to the lungs by inhalation. Many suchaerosols are known in the art, and the present invention is not limitedto any particular formulation. An example of such an aerosol is theProventil™ inhaler manufactured by Schering-Plough, the propellant ofwhich contains trichloromonofluoromethane, dichlorodifluoromethane andoleic acid. The concentrations of the propellant ingredients andemulsifiers are adjusted if necessary based on the enzyme mixture beingused in the treatment. The enzyme mixture can also be administered usinga nebulizer. When aerosol administration to the lungs is used, abronchodilator such as aminophylline, an antibiotic drug such as aβ-lactam (e.g. penicillin, cephalosporin) or quinolone, a DNase, aprotease inhibitor and/or an amiloride, can be combined with the enzymemixture for enhanced therapeutic efficacy.

The enzyme mixture can be administered in combination with antibioticsor other antimicrobial substances, other therapeutic proteins and/ormild abrasives. Suitable antibiotics include but are not limited totobramycin and duramycin. Other suitable antibiotics will be apparent tothose in the art once the organism producing the biofilm is determined.Therapeutic proteins useful in combination with the enzyme mixtures ofthe present invention include but are not limited to Dnases.

The foregoing embodiments of the present invention are further describedin the following Examples. However, the present invention is not limitedby the Examples, and variations will be apparent to those skilled in theart without departing from the scope of the present invention.

EXAMPLES Example 1 Effect of Sole Carbon Sources on the Production ofCarbohydrases

Media, chemicals and growth parameters. To assess the production ofcarbohydrases when using various carbon sources, 2-40 was grown inminimal media (Table 1) containing a final concentration of 0.2% of oneof the following carbon sources: agar or its degradation products(neoagarotetraose, neoagarobiose, D-galactose), alginic acid,carrageenan, carboxymethyl cellulose, colloidal chitin or itsdegradation products (chitobiose, chitotriose, N-acetyl-D-glucosamine),D-glucose (previously reported to repress the Microbulbifer 2-40 agarasesystem and other bacterial chitinase systems) (Stosz, 1994; Frändberg &Schnürer, 1994), laminarin (determined to repress chitinase systems inother bacteria) (Frändberg & Schnürer, 1994), -glucan (determined torepress other bacterial chitinase systems) (Frändberg & Schnürer, 1994),pectin (determined to induce other bacterial chitinase systems)(Frändberg & Schnürer, 1994), pullulan, starch (selected based onprevious finding that it repressed other bacterial chitinase systems)(Frändberg & Schnürer, 1994), xylose, or xylan. All complexcarbohydrates were added to the medium prior to autoclaving. All oligo-and mono-saccharides were prepared as 20% stocks in Pipes buffer, filtersterilized and added to cooled media (cooled to 45° C.). The cultureswere grown at room temperature with constant aeration, shaking at 200rpm. During bacterial growth, the OD_(600nm) was determined, comparedwith a standard viable growth curve to obtain cell counts, and growthcurves were generated. All chemicals were purchased from Sigma ChemicalCo. (St. Louis, Mo.), except for chitobiose and chitotriose (Seikagaku,Rockville, Md.) and agarose (FMC, Rockland, Me.). TABLE 1 Media. NameComposition MM (minimal medium) 23 g/L sea salts 1.0 g/L yeast extract2.0 g/L polysaccharide^(a) 50 ml/L Tris-HCL, pH 7.6 0.5 g/L NH₄Cl LM(Luria Marine)^(b) 10 g/L bacto-tryptone 5 g/L yeast extract 20 g/L NaCl0.1 g/L chitin adjust pH to 7.5 MM agar plates MM + 1.5% agar Chitin MMplates 23 g/L sea salts 1.0 g/L yeast extract 2.0 g/L chitin paste^(c)20 g/L phytagel MA plates (Marine agar) 37.4 g/L Marine broth 1.5 g/Lagar^(a)Insoluble polysaccharides were added to media prior to autoclaving.Other carbon sources were filter sterilized in 20 mM Pipes buffer, pH6.8 and added to cooled media to produce a final concentration of 0.2%.^(b)Vibrio harveyi BB7-1 was cultured in this medium^(c)chitin paste was produced as outlined by Lingappa and Lockwood(1962).

TABLE 2 Reagents. Reagent Composition Silver Stain Fixing SolutionEtOH:glacial acetic acid:H₂O 30:10:60 Silver Stain Developer 2.5% sodiumcarbonate 0.02% formaldehyde Coomassie Blue Stain 0.125% Coomassie BlueR-250 50% Methanol 10% glacial acetic acid Coomassie Blue DestainSolution I 50% methanol 10% acetic acid Coomassie Blue Destain SolutionII 7% acetic acid 5% methanol Imidazole-Zinc Equilibration Solution 0.2M Imidazole Imidazole-Zinc Staining Solution 0.3 M ZnSO₄ Imidazole-ZincDestaining Solution 2% citric acid India Ink Stain 0.3% Tween 20/PBS0.1% India Ink LPS Stain Fixing Solution 40% EtOH 5% glacial acetic acidPeriodic Acid Oxidizing Solution 0.833% periodic acid LPS Stain Solution0.075% NaOH 0.8% silver nitrate 1.4% (v/v) ammonium hydroxide LPS StainDeveloping Solution 50 mg/L citric acid 190 μl/L formaldehyde IodineStain 0.1 M KI 0.05 M I Triton Solution 2.5% Triton X-100 in 20 mM Pipesbuffer, pH 6.8 Dinitrosalicyclic acid Reagent (DNSA) 2.14% NaOH 0.63%3,5-dinitrosalicyclic acid 0.5% phenol

TABLE 3 Buffers. Buffer Composition Pipes Buffer 20 mM Pipes buffer, pH6.8 PBS Buffer 8 g/L NaCl 0.2 g/L KCl 1.44 g/L Na₂HPO₄ 0.24 g/L KH₂PO₄adjust pH to 7.4 with HCl Immunoblotting Buffer 2.93 g/L glycine 5.81g/L Tris 0.375 g/L SDS 20% (v/v) methanol Sodium Acetate Buffer 14.8 mlof 0.2 M acetic acid 35.2 ml of 0.2 M sodium acetate 50 ml of dH₂O pH5.0 Western Development Buffer I 80 mM Tris-HCl, pH 8.0 0.4 mg/ml4-chloro-1-napthol 0.1 μg/ml H₂O₂ (for HRP labeled antibody detection)Western Development Buffer II 0.15 mg/ml 5-bromo-4-chloro-3-indolylphosphate 0.3 mg/ml nitro blue tetrazolium 100 mM Tris buffer 5mM MgCl₂ (for AP labeled antibody detection) ELISA Development Buffer24.3 mM citric acid, pH 5.0 51.4 mM NaH₂PO₄ 0.04% o-phenylenediamine0.04% H₂O₂ Carbonate Buffer 10.6 g/L Na₂CO₃ adjust to pH 9.6 AcetateBuffer 126.5 ml of 0.2 M acetic acid 373.5 ml of 0.2 M sodium acetate500 ml dH₂O Adjust pH to 5.1 Embedding fixation buffer 3.5% NaCl 3%formaldehyde 0.1% gluteraldehyde in 20 mM phosphate buffer, pH 7.0Native PAGE Treatment buffer 12.5 mM Tris, pH 6.8 10% glycerol 0.05%bromophenol blue SDS-PAGE Treatment buffer Native PAGE treatment bufferwith 2% sodium dodecyl sulfate (SDS)

Viable plate counts. For each time point viable plate counts and opticaldensity (OD₆₀₀) were made in triplicate. The culture was vortexedthoroughly, to disrupt aggregated or substrate-bound cells, and platedon MM containing the sole carbon source. Plates, depending on the typeof MM, were incubated for 24 to 48 hours.

Enzyme harvesting. Carbohydrase activity was determined in crude enzymepreparations. These preparations consisted of whole cells orsupernatant. At each time point, 100 ml of culture was centrifuged(10,000×g, 15 min., 4° C.) and the supernatant and cell pellet wereseparated. The supernatant was stored at −20° C. until used. The wholecells were washed twice in 50 ml of Pipes buffer and then resuspended in2 ml of buffer. The concentrated whole cells were also stored at −20° C.until enzyme activity was assayed.

Dinitrosalicyclic acid (DNSA) reducing sugar assay. The DNSA assay usesdinitrosalicyclic acid reagent, developed by Sumner and coworkers(Sumner & Sisler, 1944), to quantitate the amount of carbohydraseactivity (μg/ml) by measuring the resulting reducing sugars present inthe sample. In general, the 3,5-dinitrosalicyclic acid is reduced to3-amino-5-nitrosalicyclic acid and the aldehyde groups are oxidized tocarboxyl groups (Hostettler et al., 1951). Color change in DNSA reagentis detected spectrophotometrically as it becomes reduced by any reducingsugar present in a reaction mixture.

The enzyme preparation (spent media, whole cells, or concentrated enzymepreparations) (0.3 ml) was incubated with 0.7 ml of substrate (thevarious carbohydrates listed above). Substrates were prepared as 0.5%stocks except for agarose, which was 0.2%, in buffer of either pH 5.0(0.025M sodium citrate buffer) or pH 7.0 (0.01 M potassium phosphatebuffer) depending on the polysaccharide. Carboxymethyl cellulose(Pettersson & Porath, 1966), chitin (Jeuniaux, 1966), laminarin (Ruse &Mandels, 1966) and pectin (Albersheim, 1966) were prepared in pH 5.0buffer. Agarose, alginic acid (Preiss, 1966), carrageenan, pectin,pullulan, starch, and xylan were prepared in buffer of pH 7.0. Agarosealginic acid, chitin and carrageenan were boiled for 5 min. to dissolvethem in the respective buffer prior to their addition to the reactionmixture. The reaction incubation time and temperature were alsodependent upon substrate. Agarase, alginase, and xylanse activityreactions were incubated for 1 hour at 25° C., while cellulase,chitinase, carrageenase, laminarinase, pectinase, pullulanase andamylase reactions were incubated for 2 days at 30° C.

Following incubation, 1 ml of DNSA reagent (2.14% NaOH, 0.63%3,5-dinitrosalicyclic acid, 0.5% phenol) was added to the reactionmixture and the samples were boiled for 5 min. in a hot water bath.Samples were cooled to room temperature and the absorbance at 575 nm wasdetermined. The spectrophotometer was blanked against reaction mixtureswith buffer replacing the enzyme preparation. Negative controlscontained heat inactivated enzyme preparations, autoclaved prior to theaddition to the reaction mixture. The amount of reducing sugar generatedwas determined by comparison to a galactose standard curve (20-600 μggalactose or reducing sugar equivalents) following any necessaryadjustment for residual reducing sugar present in any negative controls.A standard curve was generated for each new batch of DNSA reagentprepared. DNSA assay values are recorded as μg of reducing sugarequivalents generated per ml of sample. Triplicate samples were preparedfor each reaction and their average was taken to determine the μg/mlreducing sugar produced.

Carbohydrase activity in sole carbon sources. To assess the regulationof the carbohydrases by sole growth substrate, 2-40 was grown in minimalmedium containing a final concentration of 0.2% of one of 16 sole carbonsources. Monosaccharides included: glucose, D-galactose, glucosamine,N-acetyl-D-glucosamine (NAG), and xylose. Insoluble complexpolysaccharides included: agarose, alginic acid, carrageenan,carboxymethyl cellulose (CMC), chitin, glucan, laminarin, pectin,pullulan, starch, and xylan. Batch culture growth (OD_(600nm)) wasmonitored (FIGS. 1, 2, 3 & 4) and carbohydrase activity was assayed inboth cellular and supernatant culture fractions (Tables 4 & 5).

Enzymatic activity was reported either as total relative μg/ml ofcarbohydrase activity to report 2-40 carbohydrase activity, or as units(μg/ml carbohydrase activity per pg/ml total sample protein per DNSAassay reaction time.

Example 2 Production and Purification of Enzyme Systems

Chemicals, media and bacterial growth conditions. Pseudomonas atlanticaagarase (Sigma Chemical Co., St. Louis, Mo.) and chitinase, harvestedfrom Vibrio harveyi, served as positive controls in zymograms. Brothmedia was prepared as described in Example 1 (Table 1). Cultures werealso grown on solid media. Solid agar plates were made by adding 1% agarto the MM broth recipe (Table 1).

To induce chitinase production without agarase production, 2-40 wascultured on MM plates containing a purified chitin paste and werehardened with phytagel (Table 2.1). Chitin paste was purified fromcommercial chitin as outlined by Liggappa and Lockwood (1962). Practicalgrade chitin was soaked in 1 M NaOH for 24 hours. After the chitin waswashed with dH₂O, it was soaked in 1 M HCl for 24 hours, washed againwith dH₂O, and transferred to 1 M NaOH. The alternate base/acid soakingstep was repeated four times as described. Following the final washing,the chitin was washed 4 times with 95% EtOH and dissolved in 2 vol. of12N HCl, with constant stirring for 20 min. at room temperature. Afterfiltering the solution through glass wool into an equal volume of icecold dH₂O with constant stirring, the mixture was sedimented overnight.The sediment was washed 4 times with dH₂O and the pH adjusted to 7.0with 10M NaOH. Following centrifugation (10 min. at 4,000 rpm), thechitin paste was stored in dH₂O at 4° C. until used.

Agarase purification by ultrafiltration and ammonium sulfateprecipitation. The β-agarase I and chitinase were purified andpolyclonal antibodies were raised against them to be utilized inimmunoelectron microscopy. It was previously determined (Stosz, 1994),and confirmed in these studies, that maximal agarase was produced instationary phase supernatant when the organism was grown in agarose MM.The stationary phase supernatant (28 hours of culture growth) from 4L ofculture of Microbulbifer 2-40 grown in 0.2% agarose MM was harvested bycentrifugation (10,000×g, 15 min., 4° C.). Supernatant protein wasconcentrated to approximately 100 ml using a Minitan Manostat tangentialflow ultrafiltration system (Millipore, Piscataway, N.J.) equipped with30,000 dalton molecular weight cut off tangential flow filters. A bufferexchange was performed by passing 1 L of Pipes buffer (Table 2.3)through the system until the initial volume of 100 ml was obtained. Allsubsequent steps were performed at 4° C. The ultrafiltrate was subjectedto a 40% ammonium sulfate (AS) cut. This concentration of AS waspreviously determined to adequately precipitate the β-agarase I ofinterest (Stosz, 1994). Saturated AS (4.1M) was added dropwise to theconcentrated supernatant with constant stirring. The sample wasincubated for 1 hour to allow for equilibration before centrifugation(16,000×g, 15 min.). The resulting protein pellet was resuspended in 10ml of Pipes buffer. The crude agarase preparation was desalted byovernight dialysis against Pipes buffer. Following centrifugation(16,000×g, 15 min.), the remaining insoluble pellet was discarded andthe soluble crude agarase preparation was concentrated by centrifugationin a Centriprep-30 (30 kDa MW cut off; Amicon Inc., Beverly, Mass.).This concentrated sample is referred to as the crude agarase preparationor the 40% AS cut. Total protein concentration and enzyme activity wereassayed, as in Example 1, with the BCA protein assay and the DNSAreducing sugar assay.

Chitinase purification. It was determined that early stationary phasesupernatant contained the bulk of chitinase activity when Microbulbifer2-40 was grown in MM supplemented with 0.2% colloidal chitin (FIG. 2.4).Four liters of early stationary phase culture supernatant (40 hours)were harvested by centrifugation and concentrated with the Minitansystems described above. One liter of 10 mM Tris buffer, pH7.3 was usedto perform a buffer exchange in the Minitan system. The resulting 100 mlof crude chitinase preparation was further concentrated inCentriprep-30s. The resulting 15 ml was referred to the crude chitinasepreparation and was stored at −70° C. until used. Total proteinconcentration and enzyme activity was determined for the crude enzymepreparation.

Vibrio harveyi chitinase harvest. V. harveyi BB7-1 was obtained andchitinase produced by it was harvested to use as a control in developingchitinase enzyme assays and zymograms. The bacterium was grown in 1 L ofLM supplemented with chitin (Table 1), to induce the clone'soverproduction of chitinase, at 25° C. with shaking (100 rpm) for 3days. The sample was centrifuged (10,000×g, 15 min., 4° C.) and thesupernatant collected. The extracellular chitinase was concentrated inCentriprep-10s and a buffer exchange was performed with 10 mM Tris, pH7.3. The crude chitinase preparation was stored at −70° C. until used.

Polyacrylamide gel electrophoresis of crude enzyme preparations. Thecrude enzyme preparations were analyzed on a discontinuous SDS- ornative-PAGE as described by Laemmli (1970) using non-denaturingconditions (excluding boiling of the sample prior to electrophoresis andthe addition of β-mercaptoethanol to the 2×PAGE treatment buffer),unless otherwise indicated. Non-denaturing conditions allowed for theenzymatic activity of separated protein bands to be determined directlyin the separating gel or an overlay gel following electrophoresis. Gelsused as agarase zymograms, to detect agarase activity, had 0.1% agaroseincorporated in the separating gel. To detect chitinase activity, 0.05%glycol chitin, prepared as described below, was included in a duplicateoverlay gel. Protein samples were diluted in the appropriate 2×PAGEtreatment buffer (Table 3), either native- or SDS-, and allowed toincubate for 20 min. prior to gel loading of the samples. SDS-PAGEmolecular weight standards were included on SDS-PAGES to allow forcalculation of separated protein molecular weights (Bio-Rad, Richmond,Calif.). Following separation of the proteins by electrophoresis, theseparating gel was stained by one of the following methods or processedas a zymogram.

Zymograms (activity gels). To detect proteins possessing β-agaraseactivity in PAGE gels, 0.1% agarose was incorporated into the separatinggel. Following protein separation by electrophoresis, native PAGE gelswere placed in 100 ml of Pipes buffer and washed while shaking at roomtemperature for 10 min. The buffer was drained and replaced once. Thegel was incubated overnight at 45° C. To visually identify agarases, thegel was stained with iodine solution (Table 2) for 10 min. at 25° C.while shaking. Enzymatic protein bands appeared clear on a brownbackground. SDS-PAGE zymograms were treated as described, following two20 min. initial washings of the gel in 100 ml of 2.5% Trition X-100 inPipes buffer to remove SDS, rendering the enzymes active. The iodinestains only solidified agarose, not any oligosaccharide degradationproducts, resulting from agarase activity (Ng Yin Kin, 1972). Onlyβ-agarases (I & II), degrading polysaccharide to oligosaccharide, weresemi-quantitatively stained.

Glycol chitin was produced by acetylation of glycol chitosan (SigmaChemical Co., St. Louis, Mo.), as outlined by Trundel and Asselin(1989), and used in PAGE detection of chitinases. All procedures wereperformed at room temperature unless indicated. Glycol chitosan (10 g)was dissolved in 200 ml of 10% acetic acid by grinding the mixture in amortar. After the solution stood overnight, 900 ml of methanol wereadded slowly and the resulting solution was vacuum filtered throughWhatman No. 4 paper. To the filtrate, 15 ml of acetic anhydride wasadded while stirring. A gel was produced and allowed to stand for 30min., before cutting it into small fragments. Any liquid resulting fromthe gel fragmentation was discarded. A Waring blender (4 min. at max.speed) was used to homogenize the gel pieces which were covered withmethanol. The mixture was centrifuged (27,000×g, 15 min., 4° C.) and thepellet was resuspended in 1 vol. of methanol, rehomogenized andcentrifuged as described. The final glycol chitin pellet was resuspendedin 1 L of dH₂O+0.02% NaN₃ and homogenized. The resulting 1% (w/v) glycolchitin stock was stored at 4° C. is until used.

Chitinase zymograms, containing 0.05% glycol chitin in the separatinggel, were developed as described by Pan et al. (1991). Followingelectrophoresis, the native-PAGE was incubated for 5 min. In 150 mMsodium acetate buffer, pH 5.0. The buffer was replaced with fresh bufferand the gel was incubated for 2 hrs. at 37° C. After the buffer wasremoved, the gel was stained with 0.01% (w/v) Calcoflour White M2R in500 mM Tris-HCl, pH 8.9 for 5 min. at room temperature. Calcoflour islight sensitive, so the gel was covered for staining and the remainingsteps. The stain was removed and the gel destained overnight in dH₂O atroom temperature. Chitinase bands were detected by observation of thegel on a transilluminator and photographic documentation was made with athermal printer.

Silver staining of proteins. Silver staining of electrophoreticallyseparated proteins allows for the most sensitive detection of them,detecting as little as 0.1-1.0 ng of protein in a single band, and isapproximately 100- to 1000-times more sensitive than Coomassie Bluestaining. The silver ions, following staining with silver nitrate, arebound to the side chains of the proteins' amino acids and aredifferentially reduced upon development (Merril et al., 1984). Freeamines and sulfur groups are the principal reactive groups of theprotein (Freeman, 1973; Heukeshoven & Demick, 1985; Neilsen & Brown,1984).

Silver staining was performed as outlined by Sambrook et al. (1989), amodification of the original staining procedure described by Sammons etal. (1981). All procedures were performed at 25° C., the gel was gentlyshaken during incubation periods, and HPLC grade water was used.Proteins were fixed in the gel following separation by electrophoresisby overnight incubation in 300 ml silver stain fixing solution (Table2); The fixing solution was discarded and the gel incubated for 30 min.in 30% EtOH. The gel was washed with 4 changes of HPLC water, for 10min. each, and stained with 0.1% AgNO₃ for 30 min. The AgNO₃ solutionwas then discarded and the gel was washed under a stream of HPLC waterfor 40 sec. The protein bands were developed by 300 ml of silver staindeveloping solution (Table 2) for 10-30 min., until the desired visualdevelopment of the bands was achieved. Development was ceased byincubation of the gel for 10 min. in 1% acetic acid. All gels werewashed and stored at 4° C. in dH₂O.

Coomassie blue protein staining. The Coomassie Blue R-250 protein staindescribed by Diezel et al. (1972) was used to detect predominantproteins in preparative gel when purity and trace protein detection wasnot of significant importance. Coomassie blue stain (Table 2) was heatedto 45° C. and incubated with the gel at 25° C. for 2 hours with gentleshaking. After discarding the staining solution, the gel was destainedwith Coomassie blue destaining solution (Table 2) at 25° C. withconstant shaking and several changes until protein bands weredistinguishable in the gel. If stained gel sections were used toidentify enzyme bands to be excised from a preparative gel, the gelfragments were shrunken (in methanol) or swollen (in dH₂O) until the gelfragments matched the size of the initial separating gel, just prior toexcising of the protein band. The stained outer gel fragments werematched up with the mid-section of the separating gel to estimate theposition of the enzymatic band to be excised from the gel. The excisedgel band was washed with PBS and crushed as described below. Thisstaining method was only used for native PAGE, since imidazole-zincstaining is only compatible with SDS-PAGEs.

Imidazole-zinc protein staining. The imidazole-zinc protein stainallowed for visual detection of the protein band of interest onpreparative SDS-PAGE. The protein could then be excised and destained.This method, described by Fernandez-Patron et al. (1992), allows fordetection of proteins by negatively staining them and is only slightlyless sensitive that silver staining. Following electrophoreticseparation of the proteins, the SDS-PAGE was soaked in dH₂O for 10seconds. The gel was incubated in 200 ml of 0.2M imidazole for 10 min.at 25° C. while gently shaking. The imidazole solution was removed andthe gel was negatively stained for 2 min. in 200 ml of 0.3M ZnSO₄ atroom temperature while rocking. The protein band of interest, a clearband against an opaque background, was excised form the gel anddestained for 10 min. in 2% citric acid. The gel fragment was washedwith several changes of Pipes buffer+2.5% Triton X-100. The gelfragment, containing the enzyme of interest, was washed with PBS andfinely sliced. The gel fragments were loaded into a syringe with PBS andcrushed by passing the mixture back and forth between two glass syringesconnected by an 18 gauge hub. This mixture of crushed acrylamide and PBSwas frozen at −70° C. until use.

Topographical protuberances (Degradosomes). 2-40, grown in MM containing0.2% agarose or chitin, attach to the insoluble substrate while growingon it. Additionally, both transmission and scanning electron microscopicexamination of Microbulbifer 2-40 whole cells revealed novel cellsurface structures that were elaborated coincidentally with thedegradation of agarose (FIG. 5) and chitin (FIG. 6) and the induction ofthe respective degradative enzyme system. These structures were notsynthesized by cells that were agarase and chitinase repressed byglucose. These structures are refered to generally, as degradosomes.There were several hundred degradosomes on the cell surface. They aretypically 40-60 nm wide. They extend less from the cell surface duringearly growth phases and they extend further during late culture stages.

Immunolabeling of agarase and chitinase in degradosones. To determinewhether degradative enzymes were localized in degradosomes, whole cellsand ultrathin sections of Microbulbifer 2-40 grown undercarbohydrase-inducing or -repressing conditions were immunolabeled withabsorbed anti-agarase and/or -chitinase antisera. Agarases andchitinases were concentrated and localized in the degradosomes of cellscultivated in 0.2% agarose or chitin MM, respectively. These structuresinitially appear as cell surface blebs. In later growth stages theyelongate into tubules or they form nodules. Eventually they are releasedinto the culture supernatant. These structures were not present inmid-log phase cultures of glucose grown cells which also were notlabeled with either anti-agarase or -chitinase antisera. Additionally,statistical analysis of immunolabeled thin sections provides strongevidence for the production of agarosomes and chitinosomes, as well thelack of degradosomes in controls. Control pre-immune serum did not labelwhole cells and thin sections grown in glucose, chitin or agarose MM.

Double labeling immunoelectron microscopy, using both anti-agarase and-chitinase to label the respective enzyme, was done to: 1) determine ifboth enzyme systems are induced in a single cell; 2)-when both wereproduced; 3) see whether both enzyme systems were localized in the samedegradosome. 2-40, grown in MM plus 0.2% agarose and chitin to mid-logphase, was labeled with anti-agarase antibody and anti-chitinaseantibody. Both the agarase and chitinase systems were active in cellssampled during growth. The agarase was labeled with smaller colloidalgold particles, 10 nm, and chitinase with larger particles, 20-30 nm.Double labeled whole cells and ultrathin sections showed that bothenzyme systems were synthesized in a single cell and that eachsegregated into a different degradosome.

Extracellular agarase production. Microbulbifer 2-40 synthesizes aβ-agarase system comprised of numerous extracellular agarases, withpredominant agarases of 98, 90, 60, and 42 kDa. Many of thesedegradative agarases are packaged in tubules, vesicles or otherelaborated structures. Other species, P. atlantica, Vibrio sp. strainJTO107, and a Pseudomonas-like bacteria, synthesize multiple agarases,which arguably work cooperatively to degrade the substrate (Sugano etal., 1994; Bibb et al., 1987; Belas et al., 1988; Malmqvist, 1978). Themultiple agarases of Microbulbifer 2-40 appear to be discrete enzymes,not dissociated into lower molecular weight agarases under fullyreducing conditions, determined by comparison of silver stainednative-PAGE to fully reducing SDS-PAGE and Western blots of both of gelsprobed with anti-agarases antiserum.

The addition of β-mercaptoethanol rendered the agarases inactive,suggesting that the agarase(s) have a disulfide bond essential forenzymatic activity. Boiling also resulted in inactivation. Antiserum,raised against the 98 kDa agarase, was cross reactive with thehomologous and two other Microbulbifer 2-40 agarases, suggesting thatthese enzymes share common epitopes. As determined for other agarasessynthesized by a given species, the cross reactivity of the differentMicrobulbifer 2-40 agarases with the antiserum may be attributed tocommon domains either in the substrate binding or active sites(Malmqvist, 1978). This anti-agarase antibody inhibited up to 71% ofagarase activity in partially purified preparations, suggesting that theantibody binds directly to a common domain in the agarases' active siteor elsewhere on the enzyme to sterically hinder it from degrading itssubstrate. However, Microbulbifer 2-40 agarase is not immunologicallyrelated to P. atlantica agarase, since antiserum raised against eitheragarase was not cross reactive.

Extracellular chitinase production. Microbulbifer 2-40 attaches tochitin, and agar. Such attachment is a common mechanism used by numerousmicroorganisms for ICP hydrolysis (Svitil et al., 1997; Montgomery &Kirchman, 1993; Miron & Ben-Ghedalia, 1993; Haack & Breznak, 1993). Thisis an efficient degradative mechanism, especially for marine bacteria,maintaining contact between the organism, its enzymes and the substrate,and the end products. Also, the carbohydrases would not be so vulnerableto proteolysis, “poisoning”, or dilution (Montgomery & Kirchman, 1993;Svitil et al., 1997).

Four predominant extracellular chitinases, 200, 98, 66, & 52.5 kDa, aresynthesized by Microbulbifer 2-40 when cultured in chitin MM. Asreported for Microbulbifer 2-40 agarases and other insoluble complexpolysaccharides (ICP) degradative systems, microorganisms commonlysynthesize several enzymes with like activity to degrade ICP substrates.This is also the case for bacterial production of chitinases (Harman etal., 1993; Ilyina et al., 1995; Wantabee et al., 1992 & 1990;Techkarnijanaruk et al., 1997; Bassler et al., 1991b; Vionis et al.,1996). These individual enzymes may be the result of bacterialprocessing of a single chitinase, smaller proteolytic degradativeproducts of a single genetically encoded enzyme, or they may actually beunique enzymes, each encoded by an individual gene (Techkamijanaruk etal., 1997; Wantabe et al., 1992 & 1990; Keyhani et al., 1996; Harman etal., 1993). Like agarases, Microbulbifer 2-40 chitinases appear to beindividual enzymes, not concatamers of one another. This was determinedby comparison of silver stained native-PAGE to fully-reducing SDS-PAGEand Western blots of both gels probed with anti-chitinase antibody.

The 98 kDa chitinase was selected as the antigen for polyclonal antibodyproduction. The homologous chitinase, as well as 3 immunogically relatedchitinases, were identified by the antibody. Serological crossreactivity may result from these chitinases sharing common domains forsubstrate binding or hydrolysis. This has been shown for other bacterialchitinases by sequence homology and immunological cross reactivity(Robbins et al., 1992; S. Roseman, personal communication).Additionally, the antiserum inhibited 64% of Microbulbifer 2-40chitinase activity, under experimental conditions used. Microbulbifer2-40 chitinase does not appear to share antigenically-related domainswith V. harveyi chitinase, since antiserum raised against V. harveyichitinase is not reactive against any Microbulbifer 2-40 chitinases.

An important control showed that the anti-chitinase serum did not reactwith any proteins of glucose grown cells. This confirms the chitinasezymograms, showing that chitinase activity is repressed by glucose.

Example 3 Production of Filamentous Tubules

Morphogenesis in sole and multiple carbon source MM. Microbulbifer 2-40cells grown in glucose MM have smooth surfaces during early logarithmicphase growth. Bleb-like vesicles were formed during mid-log throughstationary culture phases. Vesicles were formed due to separation of theinner and outer membrane of the cell. (These vesicles eventuallypartition from the cell body being released in late culture stages).During late culture phases in glucose MM, late stationary to deathphase, an abundance of long, filamentous tubules, coated with smallnodules, were synthesized. The tubules were −20-50 nm in diameter andtheir length extended up to several micrometers. The nodules have anapproximate diameter of 20-40 nm.

In addition to degradosomes, filamentous tubules and bleb-like vesicleswere produced during logarithmic phase growth in MM containing agar orchitin. The appearance and abundance of tubules and blebs during earlygrowth stages in ICP were similar to those produced during late culturephase in glucose MM.

A reduction in cell size was observed during growth in MM containing ICPor in late culture stages in glucose MM. Typical rod shaped cells, onaverage 1.0 μm×2.5 μm, become more stubby and more coccoid,approximately 0.5 μm×0.35 μm. The appearance of such cells correlated towith agarase or chitinase activity.

Morphogenesis of Microbulbifer 2-40 during batch growth inneoagarohexose. Microbulbifer 2-40 grown in 0.2% neoagarohexose had ageneration time of 1.5 hours and reached a maximum cell density of2.8×109 cfu/ml, following inoculation with carbohydrase-uninduced cells(glucose grown). Cell-associated agarase activity commenced at the onsetof logarithmic phase at 5 hours of growth, peaking at 9 hours.Thereafter, for the next 22 hours, cell-associated agarase activitydeclined to almost undetectable levels, while extracellular agaraseactivity steadily increased. Neoagarohexose induced agarase, 575 μg/mlreducing sugar equivalents, during log phase growth at levels consistentwith those induced by agarose (Table 4). Cells used to inoculateneoagarohexose MM were carbohydrase-uninduced, typical in appearance(rods approximately 2.0×0.5 μm on average), and their surfaces weresmooth, lacking degradosomes. However, some tubules were transfered fromwashed, glucose-grown inoculum.

By early-log phase, cell surfaces were coated with vesicles,approximately 25-100 vesicles per cell of 50-500 nm, similar to cellsgrown in ICP. Also at this stage, coccoid cells, approximately 120-180nm in diameter, formed at the poles of the rod-shaped cells. By mid- tolate-log phase, cell surfaces are coated with tubules and vesicles andthe culture contained approximately 85% coccoid cells. The rod-shapedcells released vesicles. By mid-stationary phase, the coccoid cells,coated with tubules, constituted approximately 98% of the population.Immunoprobing of tubules. To investigate tubules and whether theycontain degradative enzyme(s), antiserum was raised againstMicrobulbifer 2-40 LPS and was tested in ELISA, Eastern blotting, andLPS stained polyacrylamide gels to confirm its reactivity against theantigen. The antiserum was not cross reactive with control LPS, isolatedfrom S. typhimurium. TABLE 4 Highest relative carbohydrase activity inculture supernatants of 2-40 grown in complex and simple carbohydratesole carbon substrates^(a). Carbohydrase Activity (μg/ml reducing sugar)Carrageen- Substrate Agarase Alginase Amylase CMCase^(b) anaseChitinase^(c) Laminarinase Pectinase Pullulanse Xylanase Glucose 143 83814 31 55 16 24 0 3 8 Agarose 525 194 805 251 101 0 134 36 311 46Alginic Acid 230 633 419 20 97 0 109 0 59 50 CMC^(d) 37 27 35 352 0 0 034 0 81 Carrageenan 47 5 309 0 209 0 31 128 4 59 Chitin 244 0 823 168 0244 211 0 615 3 Galactose 46 0 15 0 0 31 63 0 27 0 Glucan 110 0 764 1650 0 268 100 714 162 Glucosamine 142 56 0 0 0 0 54 0 0 0 Laminarin 61 0589 27 0 0 1046 0 0 0 NAG^(e) 144 81 744 0 0 0 16 0 0 0 Pectin 67 18 51038 0 11 99 405 50 67 Pullulan 172 0 730 15 0 0 0 0 362 0 Starch 311 227812 113 55 0 40 0 41 0 Xylan 273 72 97 97 0 0 115 9 0 358 Xylose 285 662542 21 0 0 0 0 0 0^(a)Supernatants were harvested from 2-40 (grown in minimal mediacontaining 0.2% of the respective substrate) during late logarithmic orearly stationary phase. The fractions were assayed for the respectivecarbohydrase activity (μg/ml reducing sugar) with the DNSA reducingsugar assay.^(b)CMCase = Carboxymethyl cellulase^(c)This assay showed that chitinase was induced at low levels by chitinand at high levels by it's oligosaccharide degradation products.^(d)CMC = Carboxymethyl cellulose^(e)NAG = N-acetyl-D-glucosamine

TABLE 5 Highest relative units of carbohydrase activity in culturesupernatants of 2-40 grown in complex and simple carbohydrate solecarbon substrates^(a). Carbohydrase Activity in Units (μg reducingsugar/μg protein^(b)) Carrageen- Substrate Agarase Alginase AmylaseCMCase^(c) anase Chitinase^(d) Laminarinase Pectinase PullulanseXylanase Glucose 1.0 0.5 6.2 0.2 0.4 0.1 0.2 0 0 0 Agarose 7.2 2.7 9.03.4 1.4 0 1.5 0.4 10.0 0.5 Alginic Acid 1.4 4.1 2.6 0.1 0.6 0 0.8 0 0.30.3 CMC^(e) 0.3 0.2 0.3 2.6 0 0 0 0.3 0 0.6 Carrageenan 0.4 0 2.7 0 1.90 0.3 1.2 0 0.6 Chitin 2.2 0 5.4 1.1 0 3.6 1.4 0 4.0 0 Galactose 0 0 0 00 0 0 0 0 0 Glucan 1.3 0 10.2 2.2 0 0 3.1 1.3 9.5 0 Glucosamine 2.2 0.70 0 0 0 0.7 0 0 0 Laminarin 0.6 0 5.6 0.3 0 0 9.9 0 0 0 NAG^(f) 1.3 0.86.1 0 0 0 0.1 0 0 0 Pectin 0.4 0.1 3.0 0.2 0 0 0.6 2.4 0.1 0.4 Pullulan2.7 0 9.8 0.2 0 0 0 0 4.9 0 Starch 2.0 1.3 4.0 0.7 0.3 0 0.2 0 3.2 0Xylan 2.7 0.8 0.7 1.1 0 0 1.0 0.1 0 3.3 Xylose 1.6 3.7 2.1 0 0 0 0 0 0 0^(a)Supernatants were harvested from 2-40 (grown in minimal mediacontaining 0.2% of the respective substrate) during late logarithmic orearly stationary phase. The fractions were assayed for the respectivecarbohydrase activity (μg/ml reducing sugar) with the DNSA reducingsugar assay.^(b)Total sample protein was determined using the Pierce BCA assay.Units of activity were calculated by dividing the total enzymaticactivity (μg/ml reducing sugar) by the total sample protein (μg/mlprotein).^(c)CMCase = Carboxymethyl cellulase^(d)This assay showed that chitinase was induced at low levels by chitinand at high levels by chito-oligosaccharides.^(e)CMC = Carboxymethyl cellulose^(f)NAG = N-acetyl-D-glucosamine

Microbulbifer 2-40 whole cells, grown in 0.2% glucose or agarose MM,were labeled with the anti-LPS antibody. Tubules, produced duringmid-log phase of growth in agarose MM, were also labeled with theanti-LPS antibody. Pre-immune serum did not label the tubules in agaroseor late stage glucose cultures.

Additionally, cells grown to mid-log phase, in either agarose or chitinMM, were labeled with antiserum raised against the homologouscarbohydrase. Tubules produced during growth in agarose were labeledwith anti-agarase antibody. Cells and tubules produced during growth inchitin MM were not labeled by the anti-agarase antibody. Similarly,tubules produced during growth in chitin were labeled withanti-chitinase antibody. Cells and tubules produced during growth inagarose MM were not labeled by anti-chitinase antibody. Additionally,the circular nodules, attached to the tubules or released into thecultures, were labeled by anti-LPS, -agarase, and -chitinase in therespective cultures.

From these data, we conclude that the tubules are membraneous andcontain carbohydrases, specifically agarase or chitinase.

Production of tubules and blebs during growth in ICP or simple sugars. 2Microbulbifer 2-40 produces tubular extensions from the outer membrane.The filamentous tubules are −30-60 nm in diameter and reach lengths upto several micrometers. They are produced during late culture phases inglucose MM, during logarithmic growth in MM containing ICP orneoagarohexose as carbon source(s), and in mid- to late-log phase growthin MM containing both glucose and ICP(s). These tubules are membranousand localize agarase(s) or chitinase(s), as determined by immunoelectronmicroscopy. They appear to elongate directly from the degradosomestructures and their abundance increases with the culture duration,comprising a substantial amount of the cell mass. “Nodules” are randomon the surface of these tubules and also free in the culture. They alsopackage agarases or chitinases.

Purification of Tubules. The cell media is centrifuged at 6000 rpm for10 minutes to pellet the cells. The supernatant is then filtered throughfilter paper with a pore size of 2-10 μm. The tubules are retained bythe filter paper and thus can be removed from the surface of the filterpaper. After the tubules are purified in this manner, they can be usedwithout further purification to degrade insoluble complexpolysaccharides.

Example 4 Partial Purification of Microbulbifer 2-40 Alginase

2-40 alginases were harvested from 28 hour culture supernatant ofMicrobulbifer 2-40 grown in 0.2% alginic acid minimal media. Maximumalginase activity was found in early stationary phase spent media,consistent with other Microbulbifer 2-40 carbohydrases. The supernatantwas concentrated using the Minitan tangential flow apparatus and theenzymes precipitated with 70% ammonium sulfate. Preliminary ammoniumsulfate precipitations of 20, 30, 40, 50, 60, 70, 80% were tested and itwas found that maximum alginase activity was present in the 70%fraction. The ammonium sulfate was dialyzed out with 20 mM Pipes at 4°C., then further concentrated using tentriprep.

Zymograms. The recovered alginases were analyzed by native-PAGE 8% andalginase activity was determined by zymorgram gel overlays containing0.1% alginic acid. Following overnight incubation in 20 mM Pipes bufferat 37° C., the activity bands were visualized by staining with 0.08%toluidine blue-O in 7% glacial acetic acid, which binds the non-degradedalginic acid.

Identification of multiple alginases from Microbulbifer2-40 bynative-PAGE

-   -   (A) Silver stain of 8% native-PAGE and    -   (B) corresponding zymogram of 8% native-PA gel overlay with        molecular weights added for reference. FIG. 7, for both A and B        Lane 1, 60 μg total protein of 0.2% glucose grown 2-40; Lane 2,        40 μg total protein of partially purified concentrated alginase        preparations. Duplicate lanes of glucose Microbulbifer 2-40 cell        prep and alginase prep from the gel were. (A) silver stained        and (B) used in the zymogram overlay, then stained with        toluidine blue O. There are eight bands with alginase activity        with approximate molecular weight of 87, 66, 43, 39, 35, 27, 25        and 23 kD.

Example 5 Degradation of Pseudomonas Biofilms

Strains:

-   -   a) Microbulbifer 2-40 b) Pseudomonas aeruginosa 2-40 FRD1        (Mucoid cystic fibrosis isolate)        -   Pseudomonas aeruginosa FRD462 (polymannuronicacid producing            mutant) (J. Bact 172:2894-2900).

Biomass. One liter of Microbulbifer 2-40 was grown to late log phase(10⁹ cells ml⁻¹), harvested, brought up in 2% instant ocean (IO) andseeded onto moist biofilms of Pseudomonas aeruginosa FRD1 & 462. (Grownon nutrient broth+0.5% yeast extract with the spent medium decanted.)Microbulbifer 2-40 was incubated at 30° C. with biofilms over a timecourse of seven days.

Biofilm degradation. This was monitored visually by examination of thefilm and by the elaboration of reducing sugars (glucuronic andmannuronic acids). Following the procedures of the above examples.Background counts of biofilms of Pseudomonas aeruginosa that were notexposed to Microbulbifer 2-40 were subtracted from the experimentalsamples. The results are shown in the following table. TABLE 6 P.aeruginosa Days & μg reducing sugar ml⁻¹ strain 0 1 3 7 FRD1 0 11 48 192FRD462 0  6 70 101

FDR1, & to a lesser extent FRD462, biofilms were visibly less obviousafter 7 days when exposed to induced Microbulbifer2-40 then when exposedto sterile 10.

Example 6 Production of S. mutans Biofilm Degrading Enzyme Mixtures

Strains

-   -   a) Microbulbifer 2-40    -   b)Streptococcus mutans ATTC 25175 (type strain) Biomass.        Microbulbifer2-40 cells are grown in broth to late log phase        (10⁹ cells ml⁻¹), harvested, brought up in 2% instant ocean (10)        and seeded onto moist biofilms of Streptococcus mutans, (S.        mutans is grown on trypticase soy agar with 5% defibrinated        sheep blood at 37° C.) Microbulbifer 2-4O is incubated at 30° C.        with the biofilms in time course.

Biofilm degradation. Biofilm degradation is monitored visually byexamination of the film and by the elaboration of reducing sugars.Following the procedures of the above examples, biofilms ofStreptococcus mutans that are not exposed to Microbulibifer 2-40 and theenzymes it produces are utilized as control materials.

1. Bacterial strain Microbulbifer 2-40.
 2. An isolated structure fromMicrobulbifer 2-40 comprising at least one degradative enzyme.
 3. Theisolated structure according to claim 2, wherein the at least onedegradative enzyme degrades a polysaccharide selected from alginate,araban, carrageenan, cellulose, carboxymethylcellulose, chitin,glycogen, β-glucan, pectin, laminarin, pullulan, starch, xylan, or agar.4. The isolated structure according to claim 3, wherein thepolysaccharide is alginate.
 5. The isolated structure according to claim3, wherein the polysaccharide is cellulose.
 6. The isolated structureaccording to claim 3, wherein polysaccharide is chitin.
 7. The isolatedstructure according to claim 3, wherein the polysaccharide is agar. 8.The isolated structure according to claim 2, wherein the at least onedegradative enzyme comprises at least two degradative enzymes.
 9. Theisolated structure according to claim 8, wherein the at least twodegradative enzymes are an agarase and a chitinase.
 10. The isolatedstructure of claim 2, obtained from a process comprising: 1) growingMicrobulbifer 2-40 to mid-log phase in a medium comprising apolysaccharide; 2) centrifuging the medium to obtain a supernatant; 3)filtering the supernatant; and 4) purifying the tubules.
 11. Theisolated structure according to claim 10, wherein the polysaccharide isalginate, araban, carrageenan, cellulose, carboxymethylcellulose,chitin, glycogen, β-glucan, pectin, laminarin, pullulan, starch, xylan,or agar.
 12. The isolated structure according to claim 10, wherein themedium is centrifuged at about 6,000 rpm for about 10 minutes.
 13. Theisolated structure according to claim 10, wherein the supernatant isfiltered through a filter paper with a pore size of about 2 μm to about10 μm.
 14. The isolated structure according to claim 2, wherein theisolated structure is an isolated degradosome, an isolated vesicle, anisolated tubule, or an isolated elaborated structure.
 15. The isolatedstructure according to claim 2, wherein the isolated structure is anisolated tubule.
 16. A degradative enzyme associated with an isolatedstructure of Microbulbifer 2-40.
 17. The degradative enzyme of claim 16,wherein the enzyme degrades a polysaccharide selected from alginate,araban, carrageenan, cellulose, carboxymethylcellulose, chitin,glycogen, β-glucan, pectin, laminarin, pullulan, starch, xylan, or agar.18. The degradative enzyme of claim 17, wherein the polysaccharide isalginate, araban, carrageenan, cellulose, carboxymethylcellulose,chitin, glycogen, β-glucan, pectin, laminarin, pullulan, starch, xylan,or agar.
 19. The degradative enzyme of claim 17, wherein thepolysaccharide is alginate.
 20. The degradative enzyme of claim 17,wherein the polysaccharide is cellulose.
 21. The degradative enzyme ofclaim 17, wherein the polysaccharide is chitin.
 22. The degradativeenzyme of claim 17, wherein the polysaccharide is agar.